Polymer Brushes on Silica Nanostructures Prepared by Aminopropylsilatrane Click Chemistry: Superior Antifouling and Biofunctionality

In nanobiotechnology, the importance of controlling interactions between biological molecules and surfaces is paramount. In recent years, many devices based on nanostructured silicon materials have been presented, such as nanopores and nanochannels. However, there is still a clear lack of simple, reliable, and efficient protocols for preventing and controlling biomolecule adsorption in such structures. In this work, we show a simple method for passivation or selective biofunctionalization of silica, without the need for polymerization reactions or vapor-phase deposition. The surface is simply exposed stepwise to three different chemicals over the course of ∼1 h. First, the use of aminopropylsilatrane is used to create a monolayer of amines, yielding more uniform layers than conventional silanization protocols. Second, a cross-linker layer and click chemistry are used to make the surface reactive toward thiols. In the third step, thick and dense poly(ethylene glycol) brushes are prepared by a grafting-to approach. The modified surfaces are shown to be superior to existing options for silica modification, exhibiting ultralow fouling (a few ng/cm2) after exposure to crude serum. In addition, by including a fraction of biotinylated polymer end groups, the surface can be functionalized further. We show that avidin can be detected label-free from a serum solution with a selectivity (compared to nonspecific binding) of more than 98% without the need for a reference channel. Furthermore, we show that our method can passivate the interior of 150 nm × 100 nm nanochannels in silica, showing complete elimination of adsorption of a sticky fluorescent protein. Additionally, our method is shown to be compatible with modifications of solid-state nanopores in 20 nm thin silicon nitride membranes and reduces the noise in the ion current. We consider these findings highly important for the broad field of nanobiotechnology, and we believe that our method will be very useful for a great variety of surface-based sensors and analytical devices.

Sample cleaning: SPR and QCMD sensor surfaces were first cleaned by rinsing with a water stream, followed by ultrasonication in acetone for 5 min, sonication in isopropanol for 5 min, drying using a gas stream of N2 and finally UV O3 treatment for at least 20 min (Compact UV-Ozone Cleaner from Cyky). An additional immersion in 99.5 % EtOH with N2 drying was performed for plain gold SPR sensors to reduce Au-OH groups after UV O3 treatment. S2 Nanochannels were cleaned with 0.2 µm syringe filtered 1 % (v/v) Hellmanex. Membranes for nanopore formation were cleaned with piranha (conc. H2SO4 and 30 % H2O2 mixed in volume ratio 3:1 for 20 min).
Silanization: The methodology is described throughout the text and in Figure 1. For the ex situ method, a droplet of APS (115 µM in 99.5 % EtOH) was placed on the surface for 1 min. For the in situ method, the solution was injected into the liquid cells and the incubation time was longer (~5 min). When functionalizing nanostructures, additional rinsing in 95% ethanol was done after binding.
Crosslinker binding: For the ex situ method, sulfo-SMCC at 0.5 g/L in 10× diluted PBS buffer was placed on the sample for 1 min. For the in situ method, the solution was introduced by steady flow using concentrations in the range 0.5-2 g/L and the incubation time was longer (~15 min).
Polymer grafting: The protocol was similar to that previously used for modifying gold. S3 For ex situ modification, the silica modified with APS and sulfo-SMCC was placed inside a petri dish containing 2 or 20 kg/mol thiol-PEG dissolved at 1 g/L or 0.12 g/L in 0.9 M Na2SO4 and incubated for 1 or 2 h respectively on a shaker table set to 100 rpm at room temperature. A liquid stream of water was used to rinse the sample, followed by drying in a stream of N2. For in situ modification, the same PEG solutions were injected into the different liquid cells.

S4
AFM measurements: A NTEGRA AFM (NT-MDT) instrument was used with Tap300Al-G tips (BudgetSensors®) with 200-400 kHz resonant frequency, 20-75 N/m force constant and 10 nm tip radius. The tapping mode AFM images were recorded. Post-processing of data includes levelling using a mean plane subtraction, followed by row alignment using a second order polynomial and offsetting to zero at the lowest measured height.
XPS measurements: Borosilicate cover glass slides were used as substrates. The APS sample was prepared by incubating in 460 µM APS in 99.5 % EtOH for 5 min, followed by rinsing in 95 % EtOH (no curing) and immersion in 10× diluted PBS for 5 min before a final water rinse and N2 drying step. The same procedure was followed for the sample with APS and sulfo-SMCC with incubation in 10× diluted PBS at 1 g/L for 30 min. The XPS measurements were conducted at a PHI 5000 VersaProbe III Scanning XPS Microprobe instrument employing a monochromated Al K X-ray source (1486.7 eV). The spectra were measured with a beam size of 200 μm 2 at 50 W and 15 keV. Carbon 1s spectra were measured first. Charging of the samples was compensated by an electron gun and an Ar + ion source. The binding energy was calibrated for each sample afterwards by shifting the binding energy range such that the leading C 1s peak appears at 285 eV, as in previous studies of similar films. S4-S7 The intensity of the spectra was normalized to the low binding energy side of the background. The spectra were deconvoluted by Voigt profiles for which the Lorentzian and Gaussian widths are individual parameters. The Lorentzian width was fixed to a literature value for the specific core hole. S8 Prior to fitting a Shirley type background was subtracted.
SPR measurements: A Bionavis 220A Navi multiparameter SPR instrument was used. The diode wavelength was 670 nm. For dry films, the reflectivity was measured with a single scan between 39.2° and 50.0° (duration 2.19 s), while continuous scanning between 58.0° and 77.9° (duration 3.88 s) was used in liquid. Layer thickness for dried films as well as exclusion heights were determined with non-linear least-square fitted Fresnel models implemented with the transfermatrix method using custom MATLAB code. S9-S11 Each SPR sensor was measured prior to further functionalisation to obtain an individual Fresnel model background, thus accounting for any initial SiO2 thickness variations between different SPR sensors. All liquid injections were performed at a flow rate of 20 µL/min with a temperature set to 25 °C.
QCMD measurements: A Q-Sense E4 instrument (Biolin Scientific) was used for the QCMD measurements, and a NE-1000 syringe pump (New Era pump systems) together with an Idex V-Supporting Information S5 451 manual injection valve (Genetec) were used for flow control and purging air bubbles. The temperature was set to 25 °C and the flow rate was 150 μL/min. Data analysis and modelling was performed with Qtools, as described previously 9 using overtones 3, 5, 7, 9 and 11. The density of the film was assumed to be 1000 kg/m 3 . Overtone three was excluded in some cases when modelling the viscoelastic thickness with the Voigt model S12 to acquire a better overall fit. The shear modulus was assumed to be independent of frequency.
Fluorescence microscopy of nanochannels: Further details and fabrication procedures for the nanofluidic devices used in this work is described elsewhere. S13 Briefly, the device contains microand nanochannels made in silicon dioxide that are sealed with borosilicate glass (Si-Mat, Germany) using high-temperature bonding. Liquid was injected by loading 15 µL in reservoirs and applying 400 mbar with N2. An inverted microscope (Zeiss AxioObserver.Z1) equipped with a 63× oil immersion objective, EMCCD camera (Photometrics Evolve) and a 475 nm LED light source (Colibri 7, Zeiss) were used for imaging. Images (512×512 pixels) were recorded at 126 gain and 100 ms exposure time. Flow was pressure driven.
Ion current recording on nanopores: Silicon nitride membranes (40 × 40 µm 2 ) were prepared in 1 cm 2 Si chips following standard protocols. S14-S16 Controlled dielectric breakdown S17 was used to form a pore in a 1 M KCl electrolyte using a commercial system (Spark-E2, Northern Nanopore Instruments). The pores were "conditioned" (grown in size) in 3.6 M LiCl. Conductance was measured by an Axopatch 200B (Molecular Devices) in 1 M KCl. To record the current baseline, a steady potential of 0.1 V was applied at sampling rate 25 kHz and bandwidth of 100 kHz. Scheme S1 Synthesis of APS; 1 H NMR spectra shown.

Non-monolayer formation of APS and SMCC (ex situ data)
The ex situ method was optimized for creating monolayers of both APS and sulfo-SMCC with the specified concentrations, solvents and incubation times. For the in situ method, one needs to be somewhat careful with concentrations and incubations times to get as close to monolayers as possible, and introduce sulfo-SMCC reasonably fast after the APS. In this section we show some data on what happens if the protocols are not followed. The main purpose is to give an idea of how precisely they need to be obeyed. As will be shown, there is quite some tolerance with respect to most factors. These supplementary results can also be useful for further understanding the chemical reactions on the surface.
We first consider the APS and sulfo-SMCC binding. Using higher concentrations and longer incubation times generally resulted in thicker films (more than a monolayer). Figure S1 shows examples of dry layer thickness determined by SPR. The final PEG layer thickness is also included, but concentration and incubation time were not altered for this step. Although the data set does not present a detailed investigation on effects from concentration and incubation time, it is clear that thicker layers of APS and sulfo-SMCC are generally formed when these parameters are increased.
Additionally, the experimental variation is much higher for all layers (including PEG) compared to when the protocol is followed (in which case the variation is ± 0.1 nm). We also noted that while most of the PEG brushes were still antifouling, some of the samples that were not prepared according to protocol did not exhibit this feature.
Supporting Information S8 Figure S1 Measured dry thickness of each layer when the concentration and/or incubation time of APS and sulfo-SMCC has been increased. For each molecular weight of PEG, the samples to the left of the dashed line were prepared according to the ex situ protocol, while the concentration and/or incubation time was increased for the others. Green check box means that the PEG brush fully repelled BSA (no binding detected), red cross means that it did not.
The effect of incubation time for APS in ethanol (99.4%) or water at different concentrations was further investigated ( Figure S2). At 460 µM in ethanol there is a tendency of forming multilayers already after ~1 min. We believe this is because APS precipitates on the surface since it is close to its solubility limit at these conditions, as supported by other observations. For instance, we noted that APS dissolved in ethanol at 460 µM over time produced a turbid solution if stored at 4 °C. In comparison, lowering the concentration to 115 µM (i.e. according to protocol) gave monolayer thickness ( Figure S2). Furthermore, incubation in water gives less than a monolayer on the surface even after 1 h, showing the importance of having the right solvent.
Supporting Information One reason why sulfo-SMCC can give multilayers at increased concentrations may be the formation of micellar structures. Figure S3 shows the surface tension as a function of total sulfo-SMCC concentration, suggesting amphiphilic properties. We could, however, not investigate micelle formation in this manner due to the solubility limit of sulfo-SMCC at 5 g/L. A OneAttension Theta goniometer from Biolin Scientific was used to measure surface tension using pendant drop method. Surface tension was recorded at 15 fps for 10 s (values are time-averaged).
Before starting measurements, the glass syringe and needle were cleaned in 1% SDS and water.
The surface tension of a reference droplet of water in room temperature was found to be 72.6-72.7 mN/m. All sample surfaces were dried with a gas stream of N2 immediately prior to contact angle measurements. Measurements were performed in the order of low to high concentrations.

Figure S3
Surface tension of solutions of sulfo-SMCC in 10× diluted PBS.
Besides forming more than a monolayer, the APS layer can also slowly hydrolyse to submonolayer coverage unless cured and/or protected by sulfo-SMCC. Figure S4 shows how the APS thickness decreases when incubating different samples in water. Interestingly, already for the initial thickness we noted an influence from the solvent type used to briefly rinse the sample after APS incubation. The lower the polarity of the rinsing solvent, the more APS remains, indicating an adsorption mechanism via hydrogen-bonding or charge interactions that may be overcome by a polar solvent. If water is used for the rinsing step, the initial layer thickness is already much below monolayer ( Figure S4). A similarly strong desorption effect in water has been observed previously for APTES. S18 The curing improves the stability of the layer, in line with previous observations for silanes S18 and can likely be inferred from the increased rate of siloxane bond formation induced by heat, replacing the initial non-covalent adsorption. S19-S21 Note that as explained in the main text, curing is convenient but not a critical step because the subsequent sulfo-SMCC layer can also be used to stabilize the initial APS layer towards hydrolysis.
Supporting Information S11 Figure S4 Stability towards hydrolysis. The thickness of APS on SiO2 was measured in SPR for different treatments after APS incubation (performed according to the ex situ protocol). The samples were rinsed for ~10 s with a stream of different solvents after which some were cured. All measurements were performed after these steps. (Note that one sample was rinsed with water but not immersed in water.) Each trace corresponds to one sample surface. Note that even a quick rinse in water desorbs much of the APS if the sample has not been cured.
Additional data on in situ modification using water To confirm the importance of the solvent for the APS binding, we also tested to use water during in situ modifications monitored with SPR ( Figure S5). Although the signal from APS seems to saturate at a value which is close to the expected signal from a monolayer, much is quickly desorbed as soon as the surface is rinsed with water again, in agreement with the data in Figure   S4. This shows that water is not suitable as solvent for promoting APS binding to silica.

Figure S5
In situ modifications in SPR with water as solvent for the initial APS binding. The signal from APS is lower and the signal from sulfo-SMCC is higher. PEG binding works but the resulting surfaces are not antifouling (BSA in PBS adsorbs considerably). The arrows show the change to PBS as running buffer.

Data for APTES modification
We managed to prepare APTES layers with an average thickness very similar to APS. An example of SPR results is shown in Figure S6, where the dry thickness was determined to 0.74 nm. However, the APTES films were still more rough (see main text) and vapor phase deposition was required for reproducibility. This is especially problematic (if possible at all) when modifying, for instance, the interior of nanochannels.
Vacuum deposition of APTES was performed inside a sealed glass bell chamber connected to a DIVAC 1.4HV3C vacuum pump. While containing the cleaned silica sample, the chamber was primed by a twice repeated pump down cycle to reduce the chamber humidity and moisture content. Subsequently, a droplet of APTES was placed on a clean microscope objective glass next to the sample, following immediate evacuation to low pressure (order of a few mbar) after which the chamber was sealed from the pump with a valve and the droplet was left to evaporate for 40 min. Finally, the sample was taken out of the chamber and cured like the APS layer. Figure S6 SPR spectra before and after APTES vapor-phase functionalization. The thickness of the film is the same as obtained for APS (assuming the same RI).
Supporting Information S14

Additional XPS analysis
To confirm the click chemistry reaction between the primary amine in APS and the sulfonated NHS group in sulfo-SMCC, we performed XPS as described in the main text. Additionally, we also looked at the S 2p region to confirm that there was only trace amounts of sulphate left on the surface S22 ( Figure S7A). This confirms that the crosslinker is binding in the expected manner to the amine groups on the surface. The survey spectrum is shown in Figure S7B. Figure S7 (A) XPS spectra of the sulphate peak region for glass (black), after APS (blue) and after sulfo-SMCC (yellow). After sulfo-SMCC the S 2p signal is not significantly larger than for the other surfaces. (B) Survey spectrum.

Including the silica coating in SPR spectra modelling
To characterize the silica coating and its influence on the SPR sensor we first performed Fresnel fitting of spectra after ALD deposition ( Figure S8). Using a literature value for the (real) refractive index of SiO2 (Table S1), we fitted both thickness and extinction coefficient of the film, which depend on the SPR minimum angle and resonance width, respectively. In this step, values previously fitted for Cr and Au for bare sensors S10 were used. Each organic layer was assumed to have a certain (purely real) refractive index and a thickness was fitted subsequently after each step.
All values are summarized in Table S1.   Supporting Information S19

Physisorption of disulphides on gold
While the current work is about the development of a method for modifying silica, we consider it informative to compare with the results obtained when grafting the same PEG chains to gold as we have done in previous work. S3, S9 One observation from those studies is that the grafting density, as determined from the dry thickness after grafting, varied depending on batch and supplier of PEG even if the specified molecular weights were the same. (For instance, for 20 kg/mol grafting densities in the range 0.2-0.3 nm −2 have been obtained.) We confirmed that PEG without thiols did not physisorb to gold ( Figure S11A). Also, we noted that upon immersing samples with high PEG amounts, corresponding to grafting densities in the higher range (close to 0.3 nm −2 for 20 kg/mol) in water, there was always a downward drift in the baseline in SPR. We left the samples immersed in water for different amounts of time and measured how the dry thickness changed ( Figure S11B). Indeed, after sufficient time the PEG amount on the surface started to stabilize on values that were corresponding to grafting densities in the lower range (~0.2 nm −2 for 20 kg/mol). This shows that many chains are physisorbed, not properly end-grafted (and the "true" grafting density on gold for 20 kg/mol PEG is thus 0.2 rather than 0.3 nm −2 ). We hypothesized that the physisorbed molecules may be disulfides and indeed, introduction of TCEP eliminated the physisorption on gold (see main text).
We see two reasons why additional "dimer" PEG chains (linked by disulfides) could physisorb on the gold surface, even when it contains a PEG brush formed by the corresponding thiol-PEG "monomer". First, disulfides have a known affinity for gold just like thiols. Second, the dimeric chains will have much higher molecular weight, which means that they precipitate more easily, and the grafting solution is already tuned to be close to the cloud point. Hence, the brush and the solid surface may act as a nucleation site for precipitation of the dimers specifically.
Note that although adding TCEP may be a convenient way to only get properly end-grafted chains on gold, this chemical must not be used when grafting to silica because it interferes with the bond formation with maleimides. S24 Since the physisorption of disulfides does not occur on silica, this is not an issue in practice. However, care must be taken to ensure that the thiol-PEG used does not contain TCEP as an additive. Also, if using TCEP when grafting to gold, it should be kept in mind that the chemical degrades at high pH. S25 However, the pH will automatically be low enough (

Dry thickness [nm]
A B Figure S12 shows additional results for the in situ modification. In each case, the molecular weight of PEG is that which is not shown in the data in the main text (2 or 20 kg/mol). Overall, the data looks similar and the only noticeable difference is the altered signal from the PEG (it is higher for 20 kg/mol than for 2 kg/mol).  Black arrows indicate change from 10× diluted PBS to regular PBS.

Contact angle measurements
While all samples were fairly hydrophilic, the surface became more hydrophobic after the sulfo-SMCC modification ( Figure S14). This strengthens the view that this layer protects the APS-SiO2 bonds from water access (that leads to hydrolysis), which is why the surfaces are stable in water after sulfo-SMCC binding but not before.
A OneAttension Theta goniometer from Biolin Scientific was used to measure static contact angles (sessile drop). Contact angles were recorded 20 s after placing a water droplet on the surface.

Figure S14
Contact angles measured after the different modification steps.

Quantification of PLL-g-PEG binding and its instability
The surface coverage in mass per area can be estimated by: Here S0 is the bulk sensitivity, δ is the decay length of the evanescent field, b is the refractometric constant and ∆θ is the shift in the SPR angle. The film thickness is d. Given that d << δ, which is the case for PLL-g-PEG and the APS-SMCC-PEG (at least for 2 kg/mol PEG), the expression can be simplified: S3 Before using this expression, the parameters S0 and δ need to be estimated. We use the simulated values from Figure S9: S0 = 126 degrees per RI unit and δ = 206 nm. This gave the PLL-g-PEG coverage reported in main text based on the signal observed from binding (0.13°, Figure S15A).
Protein coverage in liquid state was calculated in the same manner but with the standard refractometric constant of b = 0.182 cm 3 /g.
We tested the stability of the PLL-g-PEG layer with respect to the surfactant SDS, which removed around half of the adsorbed amount ( Figure S15B). Using a high flow rate further removed more of the block copolymer and the surface was not protein repelling afterwards. This illustrates limitations with non-covalent grafting methods. For comparison, the APS-SMCC-PEG layers were stable when tested in the same manner.

Intensity in nanochannels during injections
For the nanostructures, an additional rinsing step with ethanol was performed to fully wash out APS from the system before introducing sulfo-SMCC. To ensure that this did not influence the APS, we checked in real-time with SPR if there was any significant desorption when washing with ethanol. Figure S17 shows that this is not the case when tested with 95% ethanol. Assuming that the other 5% is water, the results show that such small amounts of water do not lead to any hydrolysis fast enough to become a problem in practice. This means that it is feasible to thoroughly rinse whatever fluidic system is used to get rid of APS before sulfo-SMCC is introduced, to avoid that they mix in solution phase.

Figure S17
Binding of APS and rinsing in 95% ethanol monitored by SPR.
To confirm that the channels were not clogged, we also monitored the fluorescence intensity during the injections of avidin-FITC. Figure S18A shows that the protein is being transported by flow through both the microchannel and the nanochannels. The intensity is lower for the nanochannels partly due to their lower height and partly because they only occupy a fraction of the area. Still, the increase during the injection is clearly significant. As further confirmation, we could also observe a fluorescence increase in the outlet from the nanochannels during the injections. (See design in Figure S18B.) Figure S18 (A) Fluorescence intensity measured from the (passivated) channels before, during and after injection of the fluorescent protein Avidin-FITC. The error bars represent subsequent acquisitions. (B) Device design with two inlets and two outlets. The parallel nanochannels connect the two microfluidic channels.

Nanopore formation and conductance measurements
The nanopore diameter is normally estimated from the pore conductance G as: S17 Here h is the thickness of the membrane (20 nm from ellipsometry) and σ is the bulk conductivity of the electrolyte. We measured σ = 10.7 Sm −1 for the 1 M KCl solution using a CDM210 (MeterLab). Equation S3 gives a good measure of pore diameter under the assumption that that the shape is cylindrical. For the pore modified with PEG, the result should be interpreted as an effective diameter, representing a cylindrical opening containing only the electrolyte.
Fabrication by controlled dielectric breakdown is illustrated in Figure S19, where the sudden increase in current shows pore formation. To change the size of the pore, conditioning cycles were run using the protocol in the software and the change in diameter was monitored.

Figure S19
Example of controlled dielectric breakdown for nanopore formation. The pore is formed after ~330 s as evident by the sudden increase in current. The voltage is then switched off.